We acknowledge the DFG (KI-1988/5-1 to JK, NeuroCure PhD fellowship by the NeuroCure Cluster of Excellence to MLP) for funding. We also acknowledge the Imaging Core Facility of the Leibniz Research Institute for Molecular Pharmacology Berlin (FMP) for providing the imaging set up. In addition, we would like to thank Diogo Feleciano who established the Dendra2 system in the lab and provided instructions.

1. Generation of&#160;&#160;C. elegans expressing neuronal Huntingtin-Dendra2 fusion protein
<ol>
	<li>Clone the gene encoding the POI in a nematode expression vector (i.e., pPD95_75, Addgene #1494), by traditional restriction enzyme digest10, Gibson assembly11, or any method of choice. Insert a promoter to drive expression in a desired tissue or at a desired developmental stage. Insert the Dendra2 fluorophore either N- or C-terminally in frame with the POI.</li>
	<li>Generate transgenic C. elegans expressing the fusion construct (e.g., via microinjection)12. 
	NOTE: The plasmid carrying the transgene will remain as an extrachromosomal array. Integration of the construct is not necessary but can be performed if desired13. In this protocol, C. elegans were microinjected with a plasmid carrying the fusion construct huntingtin exon 1-Dendra2 (HTT-D2) under the control of the pan-neuronal promoter Prgef-1. The C. elegans expression backbone was obtained from Kreis et al.14, the huntingtin exon 1 with either Q25 or Q97 was obtained from Juenemann et al.15, and Dendra2 was obtained from Hamer et al.16.</li>
</ol>
2. Age matching and maintenance of&#160;&#160;C. elegans
<ol>
	<li>Age match all nematodes by synchronizing either with alkaline hypochlorite solution treatment17 or via egg laying for 4 h at 20 &#176;C. For egg laying, place 10 gravid adults on a freshly seeded nematode growth media (NGM) plate and leave for 4 h before removing. The eggs laid in this timespan will give rise to synchronized nematodes.</li>
	<li>Keep experimental C. elegans on nematode growth media (NGM) plates seeded with the bacterial food source E. coli OP50, following standard nematode husbandry18.</li>
	<li>Grow nematodes at 20 &#176;C to the desired stage. For this protocol, the required ages are days 4 and 10. 
	NOTE: Young adults at day 4 can be identified by the presence of eggs in their gonads and their high mobility. Aged day 10 nematodes are post-fertile, and undergo tissue deterioration and locomotive decline19.</li>
	<li>For day 10 nematodes, passage daily after the L4 stage at day 3, once nematodes are fertile, to avoid a mixed population.</li>
</ol>
3. Preparation of microscopy slides for imaging
<ol>
	<li>Prepare the microscopy slides on the day of imaging. In a microwave, melt general grade agarose at a concentration of 3% (w/v) in ddH2O. Leave to cool slightly.</li>
	<li>Cut the tip of a 1 mL pipette tip and aspirate roughly 400 &#181;L of melted agarose. Gently place a few drops of agarose onto a clean glass slide and immediately place another slide on top, making sure that a thin pad of agarose is created between the two. Let dry before gently sliding or lifting the top slide off.</li>
	<li>Place the agarose pad slides in a humidified container to prevent them from drying out. These can be used within 2-3 h. 
	NOTE: Avoid formation of small bubbles in the agarose, as the nematodes can be trapped within.</li>
</ol>
4. Definition of confocal microscope parameters
NOTE: Before mounting the nematodes and data acquisition, define all settings on the confocal acquisition software. The settings can be adapted to the imaging hardware and software of choice.
<ol>
	<li>Open the confocal software and define the laser imaging settings. Set the light path for excitation/emission for green Dendra2 at 486-553 nm and for red Dendra2 at 580-740 nm. Adjust the power and gain of both channels/lasers according to the intensity of the fluorophore. Do not change the digital gain or offset and set the pinhole as fully open.</li>
	<li>Define the acquisition setup: select a sequential channel mode and switch track every frame. Set the scan mode as frame, and the frame size as 1,024 x 1,024, with a line step of 1. Set the averaging to 2, and average by mean method and mode of unidirectional line. Set the bit depth to 8 bits.</li>
	<li>Define the multidimensional acquisitions settings for the conversion of Dendra2. For conversion and bleaching use the 405 nm diode laser set at 60% energy power. If available, activate the safe bleaching GaAsP to protect the detectors.</li>
	<li>Select a time series of two cycles, with a 0.0 ms interval in between, and normal start and stop. Start bleaching after scan 1 of 2 and repeat for 30 iterations. Stop bleaching when the intensity drops to 50%.</li>
	<li>Define the speed of acquisition/pixel dwell as fast (e.g., maximum = 12) for conversion, and medium speed (e.g., medium = 5) for capturing a snap image. 
	NOTE: The bleaching parameters defined here are guidelines. For other Dendra2 tagged POIs the laser power settings and bleaching iterations and values must be established empirically.</li>
</ol>
5. Mounting of C. elegans onto microscopy slides
NOTE: If possible, place a mounting stereomicroscope close to the confocal microscope setup and mount the nematodes just before imaging.
<ol>
	<li>On the glass cover slide on the opposite side of the agarose pad, draw a window with four squares with a permanent marker and number them.</li>
	<li>Pipette 15 &#181;L of levamisole in the middle of the agarose pad. The concentration of levamisole will vary according to the nematode&#39;s age: When imaging day 4 nematodes use 2 mM levamisole; when imaging day 10 nematodes use 0.5 mM levamisole.</li>
	<li>Transfer four nematodes into the liquid using a wire pick. With the help of an eyelash pick, gently move each individual nematode to a window square. Swivel the eyelash so that any trace of E. coli OP50 is diluted and its fluorescent background does not interfere with signal acquisition.</li>
	<li>Wait for the nematodes to almost stop moving and gently place a cover slip on top of the liquid to immobilize the nematodes in the layer of levamisole between the agarose pad and the cover slip.</li>
	<li>Place the inverted slide on the confocal stage to image the nematodes.</li>
</ol>
6. Conversion of green Dendra2: Data acquisition at time zero
NOTE: The pulse-chase experiments start by irreversibly converting the Dendra2 fusion protein from a green emitting fluorophore to a red one.
<ol>
	<li>Using the microscope&#39;s eyepiece, locate the first nematode with a 20x objective under green fluorescence. Focus on the head or tail and switch to confocal mode.</li>
	<li>Start live laser scanning with the 488 nm blue laser to visualize the green Dendra2 in the EGFP green channel (ex/em = 486-553 nm). Select a single neuron and bring it into focus. Zoom in 3x and increase the target of the laser beam4. 
	NOTE: Select one neuron per nematode. Each neuron will constitute one sample or data point.</li>
	<li>Find the maximum projection plane and, according to the brightness of the fluorophore, increase or decrease the gain or laser power to obtain a saturated but not overexposed image identifiable by the color range indicator. Once this is defined, stop the scanning. 
	NOTE: Do not irradiate the sample for too long or with too much power, as excitation with visible blue 488 nm light can also convert Dendra2, albeit slowly and less efficiently4.</li>
	<li>In the software, open the tab to select the regions of interest (ROIs) and draw a first ROI around the selected neuron. Define a larger second region of interest encompassing the nematode&#39;s head and including the first ROI.</li>
	<li>In the bleaching settings, select for the first ROI to be acquired, bleached, and analyzed. Select for the second ROI to be acquired and analyzed but not bleached.</li>
	<li>Set the speed of scanning to maximum (i.e., fast pixel dwell) and start the experiment to convert the selected Dendra2 neurons. 
	NOTE: Once the experiment is done the acquired picture will result in two images: one before and one after conversion. For the green channel, the first image should have a higher green signal that diminishes in the second image due to the conversion of the green Dendra2. For the red channel, the first image should be negative and show no signal, with a red signal appearing in the after conversion image. If the green signal does not diminish, the conversion did not occur, and the settings, such as the 405 laser power or the number of iterations, should be modified. If there is a red signal in the first image, then the 488 nm laser power used was too high and a portion of the green Dendra2 was already converted to red. In this case, a new neuron/nematode should be selected.</li>
	<li>Immediately after conversion, start live scanning with the green 561 nm laser to visualize Dendra2 in the red channel (ex/em = 580-740 nm). Find the focus and respective maximum projection of the converted neuron using the range color indicator to avoid overexposure.</li>
	<li>Quickly set the scan rate to a lower pixel dwell speed (e.g., 5x) and acquire a snapshot image of both channels at a higher resolution. This image is defined as timepoint zero (T0) after conversion. 
	NOTE: The speed of acquisition for the converted image can be varied. However, once chosen, this speed needs to stay constant throughout the data collection.</li>
	<li>Save the scan with an identifiable name and/or number, followed by the time zero label (T0). 
	NOTE: It is advisable to also save the image of the conversion experiment (step 6.6) to illustrate the lack of red signal before conversion and its appearance afterwards.</li>
</ol>
7. Imaging of converted red Dendra2 for data acquisition at a selected second time point
<ol>
	<li>To track Dendra2 degradation over time, define a second timepoint to reimage the same nematode/neuron. Select the second timepoint experimentally to address any relevant biological question. For the protocol described here, Dendra2 is imaged both at 2 h (T2) and 24 h (T24) post conversion.
	<ol>
		<li>At the selected timepoint, find the same nematode/neuron with the use of the eyepiece and red fluorescence.</li>
		<li>Open the T0 image of the respective nematode/neuron and reload/reuse the image settings. Ensure that the acquisition settings of the snapshot are precisely the same when acquiring the T0, T2, and T24 h images.</li>
		<li>Scanning live in the red channel, bring the converted red neuron into focus. Because the red Dendra2 degrades over time the range indicator will show a less intense maximum projection. Do not change any acquisition parameters and obtain a snapshot at the same speed (e.g., 5x) as the first image.</li>
	</ol>
	</li>
	<li>To track the degradation of Dendra2 after 4 h or longer, rescue the nematodes after conversion.
	<ol>
		<li>Remove the slide from the microscope immediately after converting and imaging the four nematodes. Gently remove the coverslip and with the use of a wire pick, lift each nematode from the agarose pad.</li>
		<li>Place each nematode individually on an appropriately labelled and identifiable NGM plate.</li>
		<li>For the second time point, mount the nematode again onto a fresh agarose pad and proceed with the imaging of the converted red Dendra2 following the instructions in section 6.</li>
	</ol>
	</li>
</ol>
8. Image analysis of converted Dendra2
NOTE: Analysis of the degradation of Dendra2 is performed with Fiji/ImageJ software20.
<ol>
	<li>Open Fiji and drag and drop the .lsm file into the Fiji bar. Open the T0 image taken just after conversion and the image of the same nematode taken at the selected time point after conversion (T2 or T24 h). 
	NOTE: To track the degradation of the protein of interest fused to Dendra2 only the red channel needs to be analyzed.</li>
	<li>Establish the measurement parameters from the menu: Analyze | Set Measurements. Select the Area and Integrated Density functions.</li>
	<li>Select the image obtained with the red channel. Select the Polygon Selection Tool from the Fiji bar.</li>
	<li>Identify the converted neuron on the T0 image and draw an ROI around it using the selection tool.
	<ol>
		<li>To properly identify the contours of the neuron, highlight the intensity thresholds by selecting from the bar Image | Adjust | Threshold. Drag the bar cursor to delineate the threshold and track around this area with the polygon tool. To generate an accurate ROI, it is also possible to use the contour of the selected neuron from the green channel.</li>
	</ol>
	</li>
	<li>Once the selection has been made in the red channel window, press Analyze | Measure. A pop-up window named Results will appear and include the ROI values for Area, IntDen, and RawIntDen.</li>
	<li>Perform the same process of selection and measurement for the image of the second time point (T2 or T24 h).</li>
	<li>Copy the obtained values into a spreadsheet software, taking care to appropriately record the values at T0 after conversion, and at T2 or T24 h after conversion.</li>
</ol>
9. Calculating the ratio of Dendra2 degradation
<ol>
	<li>To calculate the ratio of degradation, first assign a value of 1 (or 100%) to time the degradation from time point zero (i.e., just after conversion, when all of the red Dendra2 converted is still present). This results from dividing the value of IntRawDen of T0 by itself.</li>
	<li>To calculate the reduction of the red Dendra2 intensity signal over time, and the degradation, divide the value of RawIntDen of the second time point (e.g., T2 or T24 h) by the value of the RawIntDen of T0. The resulting number should be less than 1. These values can also be expressed as percentages, defining T0 as 100%.</li>
	<li>Repeat section 7 for each nematode converted. For a graphical representation of the degradation of Dendra2, chart a scatter plot or bar graph with the percentage or ratio values of fluorescence decrease obtained in the y axis. Apply any desired statistical analysis and illustrate it on the graph.</li>
</ol>

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The authors have nothing to disclose.

To comprehend a protein&#39;s function it is important to understand its synthesis, location, and degradation. With the development of novel, stable, and bright FPs, visualizing and monitoring POIs has become easier and more efficient. Genetically expressed fusion PAFPs such as Dendra2 are uniquely positioned to study the stability of a POI. Upon exposure to purple-blue light, Dendra2 breaks at a precise location within a triad of conserved amino acids. The fluorophore undergoes a small structural change, resulting in a ...

The proteome of a living organism is constantly renewing itself. Proteins are continuously degraded and synthesized according to the physiological demand of a cell. Some proteins are quickly eliminated, whereas others are longer lived. Monitoring protein dynamics is a simpler, more accurate, and less invasive task when using genetically encoded fluorescent proteins (FPs). FPs form autocatalytically and can be fused to any protein of interest (POI), but do not require enzymes to fold or need cofactors save for oxygen1. A newer generation of FPs has recently been engineered to switch color upon irradiation with a light pulse of determined wavelength. These photoactivatable FPs (PAFPs) allow for labeling and tracking of POIs, or the organelles or cells they reside in, and to examine their quantitative and/or qualitative parameters2. FPs make it possible to track any POI&#39;s movement, directionality, rate of locomotion, coefficient of diffusion, mobile versus immobile fractions, the time it resides in one cellular compartment, as well as its turnover rate. For specific organelles, locomotion and transport, or fission and fusion events can be determined. For a particular cell type, a cell&#39;s position, rate of division, volume, and shape can be established. Crucially, the use of PAFPs allows tracking without continuous visualization and without interference from any newly synthetized probe. Studies both in cells and in whole organisms have successfully employed PAFPs to address biological questions in vivo, such as the development of cancer and metastasis, assembly or disassembly of the cytoskeleton, and RNA-DNA/protein interactions3. In this manuscript, light microscopy and PAFPs are used to uncover the turnover rates of the aggregation-prone protein huntingtin (HTT) in vivo in a C. elegans model of neurodegenerative disease.
The protocol described here quantifies the stability and degradation of the fusion protein huntingtin-Dendra2 (HTT-D2). Dendra2 is a second-generation monomeric PAFP4 that irreversibly switches its emission/excitation spectra from green to red in response to either UV or visible blue light, with an increase in its intensity of up to 4,000-fold5,6. Huntingtin is the protein responsible for causing Huntington&#39;s disease (HD), a fatal hereditary neurodegenerative disorder. Huntingtin exon-1 contains a stretch of glutamines (CAG, Q). When the protein is expressed with over 39Q, it misfolds into a mutant, toxic, and pathogenic protein. Mutant HTT is prone to aggregation and leads to neuronal cell death and degeneration, either as&#160;short oligomeric species or as larger highly structured amyloids7.
The nematode is a model system for studying aging and neurodegeneration thanks to its ease of manipulation, isogenic nature, short lifespan, and its optical transparency8. To study the stability of HTT in vivo, a fusion construct was expressed in the nervous system of C. elegans. A HTT-D2 transgene containing either a physiological stretch of 25Qs (HTTQ25-D2) or a pathological stretch of 97Qs (HTTQ97-D2) is overexpressed pan-neuronally throughout the nematode&#39;s lifetime9. By subjecting live C. elegans to a brief and focused point of light, a single neuron is photoswitched and the converted HTT-D2 is tracked over time. To establish the amount of HTT-D2 degraded, the difference between the red signal of the freshly converted HTT-D2 is compared to the remaining red signal of HTT-D2 after a determined period of time. Therefore, it becomes possible to investigate how huntingtin is degraded when found in its expanded and toxic form compared to its physiological form; how anterior or posterior neurons respond differently to the presence of Q97 versus Q25, especially over prolonged time periods; and how the collapse of the proteostasis network (PN) during aging contribute to the differences in degradation rates. These results only describe a small set of observations on the turnover of HTT-D2. However, many more biological questions relevant to both the field of protein aggregation and proteostasis can be addressed with this in vivo application.

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in vivo quantification of protein turnover in aging c. elegans using photoconvertible dendra2

Proteins are synthesized and degraded constantly within a cell to maintain homeostasis. Being able to monitor the degradation of a protein of interest is key to understanding not only its life cycle, but also to uncover imbalances in the proteostasis network. This method shows how to track the degradation of the disease-causing protein huntingtin. Two versions of huntingtin fused to Dendra2 are expressed in the C. elegans nervous system: a physiological version or one with an expanded and pathogenic stretch of glutamines. Dendra2 is a photoconvertible fluorescent protein; upon a short ultraviolet (UV) irradiation pulse, Dendra2 switches its excitation/emission spectra from green to red. Similar to a pulse-chase experiment, the turnover of the converted red-Dendra2 can be monitored and quantified, regardless of the interference from newly synthesized green-Dendra2. Using confocal-based microscopy and due to the optical transparency of C. elegans, it is possible to monitor and quantify the degradation of huntingtin-Dendra2 in a living, aging organism. Neuronal huntingtin-Dendra2 is partially degraded soon after conversion and cleared further over time. The systems controlling degradation are deficient in the presence of mutant huntingtin and are further impaired with aging. Neuronal subtypes within the same nervous system exhibit different turnover capacities for huntingtin-Dendra2. Overall, monitoring any protein of interest fused to Dendra2 can provide important information not only on its degradation and the players of the proteostasis network involved, but also on its location, trafficking, and transport.

Two nematode strains expressing the huntingtin exon-1 protein fragment in frame with the photoconvertible protein Dendra2 were obtained via microinjection and the plasmids were kept as an extrachromosomal array. The fusion construct was expressed in the whole C. elegans nervous system from development throughout aging. Here, HTT-D2 contained either the physiological 25 polyglutamine stretch (HTTQ25-D2, Figure 1A) or a fully penetrant...

Watch this Scientific Journal Video about In Vivo Quantification of Protein Turnover in Aging C. Elegans using Photoconvertible Dendra2 at JoVE.com

In Vivo Quantification of Protein Turnover in Aging C. Elegans using Photoconvertible Dendra2

Presented here is a protocol to monitor degradation of the protein huntingtin fused to the photoconvertible fluorophore Dendra2.

In Vivo Quantification of Protein Turnover in Aging C. Elegans using Photoconvertible Dendra2

Proteins are synthesized and degraded constantly within a cell to maintain homeostasis. Being able to monitor the degradation of a protein of interest is ...

in-vivo-quantification-protein-turnover-aging-c-elegans-using

Research

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Biology

6.1K Views.  Leibniz Research Institute for Molecular Pharmacology im Forschungsverbund. Presented here is a protocol to monitor degradation of the protein huntingtin fused to the photoconvertible fluorophore Dendra2.

Video: In Vivo Quantification of Protein Turnover in Aging C. Elegans using Photoconvertible Dendra2

Generation of Stable Transgenic C. elegans Using Microinjection

Transgenic Caenorhabditis elegans can be readily created via microinjection of a DNA plasmid solution into the gonad 1. The plasmid DNA rearranges to form extrachromosomal concatamers that are stably inherited, though not with the same efficiency as actual chromosomes 2. A gene of interest is co-injected with an obvious phenotypic marker, such as rol-6 or GFP, to allow selection of transgenic animals under a dissecting microscope. The exogenous gene may be expressed from its native promoter for cellular localization studies. Alternatively, the transgene can be driven by a different tissue-specific promoter to assess the role of the gene product in that particular cell or tissue. This technique efficiently drives gene expression in all tissues of C. elegans except for the germline or early embryo 3. Creation of transgenic animals is widely utilized for a range of experimental paradigms. This video demonstrates the microinjection procedure to generate transgenic worms. Furthermore, selection and maintenance of stable transgenic C. elegans lines is described.

This video demonstrates the technique of microinjection into the gonad of C. elegans to create transgenic animals.

Transgenic Caenorhabditis elegans can be readily created via microinjection of a DNA plasmid solution into the gonad 1.  The plasmid DNA rearranges to ...

In vivo Quantification of G Protein Coupled Receptor Interactions using Spectrally Resolved Two-photon Microscopy

The study of protein interactions in living cells is an important area of research because the information accumulated both benefits industrial applications as well as increases basic fundamental biological knowledge. F&ouml;rster (Fluorescence) Resonance Energy Transfer (FRET) between a donor molecule in an electronically excited state and a nearby acceptor molecule has been frequently utilized for studies of protein-protein interactions in living cells. The proteins of interest are tagged with two different types of fluorescent probes and expressed in biological cells. The fluorescent probes are then excited, typically using laser light, and the spectral properties of the fluorescence emission emanating from the fluorescent probes is collected and analyzed. Information regarding the degree of the protein interactions is embedded in the spectral emission data. Typically, the cell must be scanned a number of times in order to accumulate enough spectral information to accurately quantify the extent of the protein interactions for each region of interest within the cell. However, the molecular composition of these regions may change during the course of the acquisition process, limiting the spatial determination of the quantitative values of the apparent FRET efficiencies to an average over entire cells. By means of a spectrally resolved two-photon microscope, we are able to obtain a full set of spectrally resolved images after only one complete excitation scan of the sample of interest. From this pixel-level spectral data, a map of FRET efficiencies throughout the cell is calculated. By applying a simple theory of FRET in oligomeric complexes to the experimentally obtained distribution of FRET efficiencies throughout the cell, a single spectrally resolved scan reveals stoichiometric and structural information about the oligomer complex under study. Here we describe the procedure of preparing biological cells (the yeast Saccharomyces cerevisiae) expressing membrane receptors (sterile 2 &alpha;-factor receptors) tagged with two different types of fluorescent probes. Furthermore, we illustrate critical factors involved in collecting fluorescence data using the spectrally resolved two-photon microscopy imaging system. The use of this protocol may be extended to study any type of protein which can be expressed in a living cell with a fluorescent marker attached to it.

In vivo Quantification of G Protein Coupled Receptor Interactions using Spectrally Resolved Two-photon Microscopy

By employing a spectrally resolved two-photon microscopy imaging system, pixel-level maps of F&ouml;rster Resonance Energy Transfer (FRET) efficiencies are obtained for cells expressing membrane receptors hypothesized to form homo-oligomeric complexes. From the FRET efficiency maps, we are able to estimate stoichiometric information about the oligomer complex under study.

The study of protein interactions in living cells is an important area of research because the information accumulated both benefits industrial ...

Monitoring of Ubiquitin-proteasome Activity in Living Cells Using a Degron (dgn)-destabilized Green Fluorescent Protein (GFP)-based Reporter Protein

Proteasome is the main intracellular organelle involved in the proteolytic degradation of abnormal, misfolded, damaged or oxidized proteins 1, 2. Maintenance of proteasome activity was implicated in many key cellular processes, like cell's stress response 3, cell cycle regulation and cellular differentiation 4 or in immune system response 5. The dysfunction of the ubiquitin-proteasome system has been related to the development of tumors and neurodegenerative diseases 4, 6. Additionally, a decrease in proteasome activity was found as a feature of cellular senescence and organismal aging 7, 8, 9, 10. Here, we present a method to measure ubiquitin-proteasome activity in living cells using a GFP-dgn fusion protein. To be able to monitor ubiquitin-proteasome activity in living primary cells, complementary DNA constructs coding for a green fluorescent protein (GFP)&#x2013;dgn fusion protein (GFP&#x2013;dgn, unstable) and a variant carrying a frameshift mutation (GFP&#x2013;dgnFS, stable 11) are inserted in lentiviral expression vectors. We prefer this technique over traditional transfection techniques because it guarantees a very high transfection efficiency independent of the cell type or the age of the donor. The difference between fluorescence displayed by the GFP&#x2013;dgnFS (stable) protein and the destabilized protein (GFP-dgn) in the absence or presence of proteasome inhibitor can be used to estimate ubiquitin-proteasome activity in each particular cell strain. These differences can be monitored by epifluorescence microscopy or can be measured by flow cytometry.

Monitoring of Ubiquitin-proteasome Activity in Living Cells Using a Degron dgn-destabilized Green Fluorescent Protein GFP-based Reporter Protein

A method to monitor ubiquitin-proteasome activity in living cells is described. A degron-destabilized GFP- (GFP-dgn) and a stable GFP-dgnFS fusion protein are generated and transduced into the cell using a lentiviral expression vector. This technique allows to generate a stable GFP-dgn/GFP-dgnFS expressing cell line in which ubiquitin-proteasome activity can be easily assessed using epifluorescence or flow cytometry.

Proteasome is the main intracellular organelle involved in the proteolytic degradation of abnormal, misfolded, damaged or oxidized proteins 1, 2. ...

Antibody Staining in C. Elegans Using &quot;Freeze-Cracking&quot;

To stain C. elegans with antibodies, the relatively impermeable cuticle must be bypassed by chemical or mechanical methods. "Freeze-cracking" is one method used to physically pull the cuticle from nematodes by compressing nematodes between two adherent slides, freezing them, and pulling the slides apart. Freeze-cracking provides a simple and rapid way to gain access to the tissues without chemical treatment and can be used with a variety of fixatives. However, it leads to the loss of many of the specimens and the required compression mechanically distorts the sample. Practice is required to maximize recovery of samples with good morphology. Freeze-cracking can be optimized for specific fixation conditions, recovery of samples, or low non-specific staining, but not for all parameters at once. For antibodies that require very hard fixation conditions and tolerate the chemical treatments needed to chemically permeabilize the cuticle, treatment of intact nematodes in solution may be preferred. If the antibody requires a lighter fix or if the optimum fixation conditions are unknown, freeze-cracking provides a very useful way to rapidly assay the antibody and can yield specific subcellular and cellular localization information for the antigen of interest.

Antibody Staining in C. Elegans Using &quot;Freeze-Cracking&quot;

"Freeze-cracking," a method for exposing the inner tissues of the nematode C. elegans to antibodies for protein localization, is demonstrated.

To stain C.  elegans with antibodies, the relatively impermeable cuticle must be  bypassed by chemical or mechanical methods.  "Freeze-cracking" is one ...

Using Caenorhabditis elegans as a Model System to Study Protein Homeostasis in a Multicellular Organism

The folding and assembly of proteins is essential for protein function, the long-term health of the cell, and longevity of the organism. Historically, the function and regulation of protein folding was studied in vitro, in isolated tissue culture cells and in unicellular organisms. Recent studies have uncovered links between protein homeostasis (proteostasis), metabolism, development, aging, and temperature-sensing. These findings have led to the development of new tools for monitoring protein folding in the model metazoan organism Caenorhabditis elegans. In our laboratory, we combine behavioral assays, imaging and biochemical approaches using temperature-sensitive or naturally occurring metastable proteins as sensors of the folding environment to monitor protein misfolding. Behavioral assays that are associated with the misfolding of a specific protein provide a simple and powerful readout for protein folding, allowing for the fast screening of genes and conditions that modulate folding. Likewise, such misfolding can be associated with protein mislocalization in the cell. Monitoring protein localization can, therefore, highlight changes in cellular folding capacity occurring in different tissues, at various stages of development and in the face of changing conditions. Finally, using biochemical tools ex vivo, we can directly monitor protein stability and conformation. Thus, by combining behavioral assays, imaging and biochemical techniques, we are able to monitor protein misfolding at the resolution of the organism, the cell, and the protein, respectively.

Using Caenorhabditis elegans as a Model System to Study Protein Homeostasis in a Multicellular Organism

To study the relationship between protein homeostasis, stress and aging, we monitored changes in protein folding by following protein dysfunction, protein localization in the cell and protein stability at the organismal, cellular and protein levels, using the genetically tractable metazoan Caenorhabditis elegans as a model system.

The folding and assembly of proteins is essential for protein function, the long-term health of the cell, and longevity of the organism. Historically, the ...

Methods to Study Changes in Inherent Protein Aggregation with Age in Caenorhabditis elegans

In the last decades, the prevalence of neurodegenerative disorders, such as Alzheimer's disease (AD) and Parkinson's disease (PD), has grown. These age-associated disorders are characterized by the appearance of protein aggregates with fibrillary structure in the brains of these patients. Exactly why normally soluble proteins undergo an aggregation process remains poorly understood. The discovery that protein aggregation is not limited to disease processes and instead part of the normal aging process enables the study of the molecular and cellular mechanisms that regulate protein aggregation, without using ectopically expressed human disease-associated proteins. Here we describe methodologies to examine inherent protein aggregation in Caenorhabditis elegans through complementary approaches. First, we examine how to grow large numbers of age-synchronized C. elegans to obtain aged animals and we present the biochemical procedures to isolate highly-insoluble-large aggregates. In combination with a targeted genetic knockdown, it is possible to dissect the role of a gene of interest in promoting or preventing age-dependent protein aggregation by using either a comprehensive analysis with quantitative mass spectrometry or a candidate-based analysis with antibodies. These findings are then confirmed by in vivo analysis with transgenic animals expressing fluorescent-tagged aggregation-prone proteins. These methods should help clarify why certain proteins are prone to aggregate with age and ultimately how to keep these proteins fully functional.

Methods to Study Changes in Inherent Protein Aggregation with Age in Caenorhabditis elegans

The goal of the method presented here is to explore protein aggregation during normal aging in the model organism C. elegans. The protocol represents a powerful tool to study the highly insoluble large aggregates that form with age and to determine how changes in proteostasis impact protein aggregation.

In the last decades, the prevalence of neurodegenerative disorders, such as Alzheimer's disease (AD) and Parkinson's disease (PD), has grown. These ...

Quantification of Site-specific Protein Lysine Acetylation and Succinylation Stoichiometry Using Data-independent Acquisition Mass Spectrometry

Post-translational modification (PTM) of protein lysine residues by N&#400;-acylation induces structural changes that can dynamically regulate protein functions, for example, by changing enzymatic activity or by mediating interactions. Precise quantification of site-specific protein acylation occupancy, or stoichiometry, is essential for understanding the functional consequences of both global low-level stoichiometry and individual high-level acylation stoichiometry of specific lysine residues. Other groups have reported measurement of lysine acetylation stoichiometry by comparing the ratio of peptide precursor isotopes from endogenous, natural abundance acylation and exogenous, heavy isotope-labeled acylation introduced after quantitative chemical acetylation of proteins using stable isotope-labeled acetic anhydride. This protocol describes an optimized approach featuring several improvements, including: (1) increased chemical acylation efficiency, (2) the ability to measure protein succinylation in addition to acetylation, and (3) improved quantitative accuracy due to reduced interferences using fragment ion quantification from data-independent acquisitions (DIA) instead of precursor ion signal from data-dependent acquisition (DDA). The use of extracted peak areas from fragment ions for quantification also uniquely enables differentiation of site-level acylation stoichiometry from proteolytic peptides containing more than one lysine residue, which is not possible using precursor ion signals for quantification. Data visualization in Skyline, an open source quantitative proteomics environment, allows for convenient data inspection and review. Together, this workflow offers unbiased, precise, and accurate quantification of site-specific lysine acetylation and succinylation occupancy of an entire proteome, which may reveal and prioritize biologically relevant acylation sites.

Here, we present unbiased quantification of site-specific protein acetylation and/or succinylation occupancy (stoichiometry) of an entire proteome through a ratiometric analysis of endogenous modifications to modifications introduced after quantitative chemical acylation using stable isotope-labeled anhydrides. In combination with sensitive data-independent acquisition mass spectrometry, accurate site occupancy measurements are obtained.

Post-translational modification (PTM) of protein lysine residues by N&#400;-acylation induces structural changes that can dynamically regulate protein ...

Measurements of Physiological Stress Responses in C. Elegans

Organisms are often exposed to fluctuating environments and changes in intracellular homeostasis, which can have detrimental effects on their proteome and physiology. Thus, organisms have evolved targeted and specific stress responses dedicated to repair damage and maintain homeostasis. These mechanisms include the unfolded protein response of the endoplasmic reticulum (UPRER), the unfolded protein response of the mitochondria (UPRMT), the heat shock response (HSR), and the oxidative stress response (OxSR). The protocols presented here describe methods to detect and characterize the activation of these pathways and their physiological consequences in the nematode, C. elegans. First, the use of pathway-specific fluorescent transcriptional reporters is described for rapid cellular characterization, drug screening, or large-scale genetic screening (e.g., RNAi or mutant libraries). In addition, complementary, robust physiological assays are described, which can be used to directly assess sensitivity of animals to specific stressors, serving as functional validation of the transcriptional reporters. Together, these methods allow for rapid characterization of the cellular and physiological effects of internal and external proteotoxic perturbations.

Measurements of Physiological Stress Responses in C. Elegans

Here, we characterize cellular proteotoxic stress responses in the nematode C. elegans by measuring the activation of fluorescent transcriptional reporters and assaying sensitivity to physiological stress.

Organisms are often exposed to fluctuating environments and changes in intracellular homeostasis, which can have detrimental effects on their proteome and ...

Measuring RAN Peptide Toxicity in C. elegans

C. elegans is commonly used to model age-related neurodegenerative diseases caused by repeat expansion mutations, such as Amyotrophic Lateral Sclerosis (ALS) and Huntington&#8217;s disease. Recently, repeat expansion-containing RNA was shown to be the substrate for a novel type of protein translation called repeat-associated non-AUG-dependent (RAN) translation. Unlike canonical translation, RAN translation does not require a start codon and only occurs when repeats exceed a threshold length. Because there is no start codon to determine the reading frame, RAN translation occurs in all reading frames from both sense and antisense RNA templates that contain a repeat expansion sequence. Therefore, RAN translation expands the number of possible disease-associated toxic peptides from one to six. Thus far, RAN translation has been documented in eight different repeat expansion-based neurodegenerative and neuromuscular diseases. In each case, deciphering which RAN products are toxic, as well as their mechanisms of toxicity, is a critical step towards understanding how these peptides contribute to disease pathophysiology. In this paper, we present strategies to measure the toxicity of RAN peptides in the model system C. elegans. First, we describe procedures for measuring RAN peptide toxicity on the growth and motility of developing C. elegans. Second, we detail an assay for measuring postdevelopmental, age-dependent effects of RAN peptides on motility. Finally, we describe a neurotoxicity assay for evaluating the effects of RAN peptides on neuron morphology. These assays provide a broad assessment of RAN peptide toxicity and may be useful for performing large-scale genetic or small molecule screens to identify disease mechanisms or therapies.

Measuring RAN Peptide Toxicity in C. elegans

Repeat-associated non-ATG-dependent translational products are emerging pathogenic features of several repeat expansion-based diseases. The goal of the protocol described is to evaluate toxicity caused by these peptides using behavioral and cellular assays in the model system C. elegans.

C. elegans is commonly used to model age-related neurodegenerative diseases caused by repeat expansion mutations, such as Amyotrophic Lateral Sclerosis ...

Label-Free Imaging of Lipid Storage Dynamics in Caenorhabditis elegans using Stimulated Raman Scattering Microscopy

Lipid metabolism is a fundamental physiological process necessary for cellular and organism health. Dysregulation of lipid metabolism often elicits obesity and many associated diseases including cardiovascular disorders, type II diabetes, and cancer. To advance the current understanding of lipid metabolic regulation, quantitative methods to precisely measure in vivo lipid storage levels in time and space have become increasingly important and useful. Traditional approaches to analyze lipid storage are semi-quantitative for microscopic assessment or lacking spatio-temporal information for biochemical measurement. Stimulated Raman scattering (SRS) microscopy is a label-free chemical imaging technology that enables rapid and quantitative detection of lipids in live cells with a subcellular resolution. As the contrast is exploited from intrinsic molecular vibrations, SRS microscopy also permits four-dimensional tracking of lipids in live animals. In the last decade, SRS microscopy has been widely used for small molecule imaging in biomedical research and overcome the major limitations of conventional fluorescent staining and lipid extraction methods. In the laboratory, we have combined SRS microscopy with the genetic and biochemical tools available to the powerful model organism, Caenorhabditis elegans, to investigate the distribution and heterogeneity of lipid droplets across different cells and tissues and ultimately to discover novel conserved signaling pathways that modulate lipid metabolism. Here, we present the working principles and the detailed setup of the SRS microscope and provide methods for its use in quantifying lipid storage at distinct developmental timepoints of wild-type and insulin signaling deficient mutant C. elegans.

Label-Free Imaging of Lipid Storage Dynamics in Caenorhabditis elegans using Stimulated Raman Scattering Microscopy

Stimulated Raman scattering (SRS) microscopy allows selective, label-free imaging of specific chemical moieties and it has been effectively employed to image lipid molecules in vivo. Here, we provide a brief introduction to the principle of SRS microscopy and describe methods for its use in imaging lipid storage in Caenorhabditis elegans.

Lipid metabolism is a fundamental physiological process necessary for cellular and organism health. Dysregulation of lipid metabolism often elicits ...